Understanding the photophysics of the Spinach-DFHBI RNA aptamer-fluorogen complex to improve live-cell RNA imaging

Article abstract of DOI:10.1021/ja411060p

The use of aptamer-fluorogen complexes is an emerging strategy for RNA imaging. Despite its promise for cellular imaging and sensing, the low fluorescence intensity of the Spinach-DFHBI RNA aptamer-fluorogen complex hampers its utility in quantitative liv

Full text of DOI:10.1021/ja411060p

Article
pubs.acs.org/JACS
Understanding the Photophysics of the Spinach−DFHBI RNA
Aptamer−Fluorogen Complex To Improve Live-Cell RNA Imaging
Kyu Young Han,†,‡ Benjamin J. Leslie,†,‡ Jingyi Fei,‡ Jichuan Zhang,§ and Taekjip Ha*,†,‡,∥
†Howard Hughes Medical Institute, Urbana, Illinois 61801, United States
‡Department of Physics and Center for the Physics of Living Cells, §Department of Materials Science and Engineering, and ∥Institute
for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
S
* Supporting Information
ABSTRACT: The use of aptamer−fluorogen complexes is an
emerging strategy for RNA imaging. Despite its promise for cellular
imaging and sensing, the low fluorescence intensity of the
Spinach−DFHBI RNA aptamer−fluorogen complex hampers its
utility in quantitative live-cell and high-resolution imaging
applications. Here we report that illumination of the Spinach−
fluorogen complex induces photoconversion and subsequently
fluorogen dissociation, leading to fast fluorescence decay and
fluorogen-concentration-dependent recovery. The fluorescence
lifetime of Spinach−DFHBI is 4.0
0.1 ns irrespective of the
extent of photoconversion. We detail a low-repetition-rate illumination scheme that enables us to maximize the potential of the
Spinach−DFHBI RNA imaging tag in living cells.
Recently, a simple and powerful method was introduced that
facilitates RNA detection. A nontoxic and membrane-
permeable nonfluorescent GFP chromophore analogue (or
fluorogen), 3,5-difluoro-4-hydroxybenzylidene imidazolinone
(DFHBI), binds to a genetically encoded RNA aptamer called
“Spinach,” eliciting green fluorescence (Figure 1a).11 Indeed,
Spinach was successfully implemented for visualization of
ribosomal RNA in living mammalian cells11 and sensing of
intracellular metabolites and protein expression by means of
modular aptamers.12 This fluorescent tag motivates potential
applications to quantitative imaging of low-abundance RNA
targets.
INTRODUCTION
■
RNA has important roles in various cellular and developmental
processes. It not only functions as a passive intermediary in
protein production based on genetic information but also is a
post-transcriptional regulator of gene expression.1 Fluorescence
imaging is one of the most valuable tools for studying these
diverse functions of RNA by enabling direct visualization of
RNA molecules in the cell and providing temporal and spatial
information.2 For example, it has been used to characterize
transcription, transport, localization, translation, and degrada-
tion of messenger RNA3 and to reveal critical roles played by
noncoding RNA.4
Under cellular imaging conditions, however, Spinach shows
greatly reduced fluorescence compared with what is expected
on the basis of its nominal brightness, which is reported to be
similar to that of GFP.11,13 Here we present a plausible
mechanism to explain its limited fluorescence intensity and
show that this aptamer−fluorogen complex has novel proper-
ties distinct from typical fluorescent proteins. Exploiting its
underlying photophysical properties, we demonstrate that >5-
fold higher total photon flux and 10-fold higher steady-state
fluorescence intensity can be obtained by low-repetition-rate
illumination in living Escherichia coli.
Several RNA-imaging techniques have been developed, of
which the most widely used are fluorescence in situ
hybridization (FISH) with organic-dye-labeled oligonucleotide
probes in fixed cells5 and fluorescent-protein-fused RNA
binding proteins in living cells.6−8 In particular, the MS2
system has been widely used for in vivo labeling of transcripts
containing repeated stem−loops, which are specifically bound
by a coat protein of bacteriophage MS2 fused to a green
fluorescent protein (GFP).6 Each of these two techniques has a
key limitation of its utility. RNA FISH has single-RNA
sensitivity but is largely limited to fixed cells. MS2 systems
can be used in live cells, but high background fluorescence from
unbound GFP limits their sensitivity. So-called “split-GFP”
approaches significantly decrease the background signal,8,9 but
at a considerable cost to the fluorescence intensity of assembled
GFP fluorophores.10 While these techniques are complemen-
tary, a single technique that combines the sensitivity of FISH
with live-cell capabilities would be valuable.
MATERIALS AND METHODS
■
Chemical Synthesis of DFHBI. DFHBI was synthesized from 4-
hydroxy-3,5-difluorobenzaldehyde as a starting reagent following the
Received: October 29, 2013
Published: November 28, 2013
© 2013 American Chemical Society
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5 min and then washed out. After incubation of GFP (5 nM) in the
flow chamber for 10 min, unbound excess proteins were flushed before
measurements were taken.
Fluorescence Intensity and Lifetime Measurements. We used
a custom-built confocal microscope to detect fluorescence of GFP and
Spinach, unless specified otherwise.16 A blue diode laser (473 nm, CNI
laser) was used to illuminate the sample, and the illumination time and
level of light intensity were controlled by acousto-optical filters
(MT200-A0.5-VIS, AA Opto-Electronic). The fluorescence lifetime
measurements were performed using the same microscope with an
ultrafast laser (MaiTai HP, Spectra Physics) and a time-correlated
single photon counting module (SPC630, Becker Hickl).17 Please refer
to the SI for more information.
Binding and Unbinding Kinetics Measurements. The
fluorescence increase was recorded with a fluorometer (Cary Eclipse)
after rapid mixing of 40 nM Spinach with different concentrations of
DFHBI in a cuvette (80 μL). The excitation and emission wavelengths
were 460 and 510 nm, respectively. The illumination intensity was
estimated to be 1−5 W/cm2. The fluorescence intensity followed a
single-exponential time course.
Live-Cell Imaging in E. coli. Rosetta (DE3) competent cells
(Novagen) were transformed with pET28c-tRNA-Spinach (provided
by Prof. Samie Jaffrey), which encodes a Spinach aptamer inserted into
the tRNALys3 sequence. Cultures were grown overnight with shaking in
Luria Broth (LB) containing 50 μg/mL kanamycin at 37 °C. Then
aliquots of the static culture were added to 5 mL of LB containing 1
mM isopropyl β-D-1-thiogalactopyranoside (IPTG) to reach an optical
density of 0.4 at 600 nm and induced at 37 °C. After 2.5 h, 100 μL
aliquots of cells were added to individual wells of an eight-well Lab-
Tek chamber (NUNC no. 155411) coated with polylysine, and cells
were allowed to adhere for 30 min at 25 °C. Cell-coated surfaces were
gently washed two times with 200 μL of modified minimal M9CA
broth (Teknova no. M8011) with glycerol supplemented with 1 mM
IPTG and then incubated for 30 min at 25 °C in 200 μL of M9CA
containing 1 mM IPTG and 100 μM DFHBI. We used a home-built
epi-illumination microscope to image live cells with an oil immersion
objective lens (NA = 1.4, UPlanSApo 100×, Olympus) and an
EMCCD camera (iXon DU-897, Andor). The illumination time of
473 nm light was synchronized to clock signals of the camera and
controlled by a custom program. Image analysis was performed using
MATLAB (The MathWorks) and ImageJ (NIH) software.
Figure 1. (a) The binding of the GFP-like fluorogen DFHBI to the
RNA aptamer Spinach activates green fluorescence upon 473 nm
illumination. (b) Fluorescence time traces of GFP (green) and
Spinach (red) immobilized on a surface and subjected to light that was
turned on and off periodically (blue, Iexc = 0.2 kW/cm2). The
Spinach−DFHBI complex shows fast fluorescence decay and
reversible recovery in the dark, whereas GFP displays irreversible
fluorescence decay on this time scale.
procedure of Paige and co-workers [see the Supporting Information
(SI) for details].11
Preparation of the Spinach RNA Aptamer. DNA coding for the
Spinach aptamer was cloned into the pET28a plasmid (Novagen).
Spinach was subsequently transcribed in vitro from the T7 promoter
using a MEGAshortscript T7 Kit (Ambion) according to the protocol
provided by the manufacturer. Then the transcribed RNA was buffer-
exchanged with a Micro Bio-Spin P-6 column (Bio-Rad) three times
into storage buffer (10 mM Tris-HCl, pH 7.5, 0.1 mM EDTA, and 10
mM KCl) to remove free, unincorporated nucleotides. The Spinach
RNA was then folded in 40 mM HEPES buffer (pH 7.5) with 125 mM
KCl by incubation at 90 °C for 2 min and slow cooling to room
temperature. At 65 °C, MgCl2 was supplemented to a final
concentration of 5 mM, and cooling was resumed.11 For experiments
requiring attachment of Spinach to surfaces, a biotin-labeled DNA
oligonucleotide (5′-TTATCCGCTCACAATTCCCATTTG-biotin-
3′, Integrated DNA Technologies) was annealed to the Spinach 5′-
end by direct mixing of the biotin−DNA with Spinach during folding
as described above.
Surface Immobilization of GFP and Spinach. A flow chamber
was prepared on a poly(ethylene glycol) (PEG)-coated coverslip with
a low density of biotin−PEG (Laysan Bio).14 Neutravidin (0.2 mg/
mL, Thermo Scientific) was incubated for 5 min and washed away
with 1× PBS buffer (Lonza no. 17-517Q). Freshly folded biotinylated
Spinach aptamer (10 nM) was incubated for 10 min in an imaging
buffer consisting of 40 mM HEPES (pH 7.5) with 125 mM KCl and 5
mM MgCl2.11 Then various concentrations of DFHBI (50 μL) in the
same buffer were loaded to the chamber for the measurements. The
surface density of the immobilized RNA was roughly 5−20 molecules
per focal volume. In the case of fluorescent proteins, recombinant
Aequorea coerulescens GFP (rAcGFP1) was purchased from Clontech
and used without further purification. The spectral properties of
rAcGFP1 (hereinafter simply GFP) are reported to be similar to those
of enhanced GFP.15 To immobilize GFP to the PEG surface, a
biotinylated rabbit polyclonal anti-GFP antibody (Rockland Immu-
nochemicals) was incubated on Neutravidin-coated flow chambers for
RESULTS AND DISCUSSION
■
Fast Fluorescence Intensity Decay and Recovery of
the Spinach−DFHBI Complex. Figure 1b shows typical
fluorescence intensity time traces of the surface-immobilized
fluorescent protein and Spinach at high-density coverage under
a focused laser beam. While blue laser light (473 nm) with an
excitation intensity (Iexc) of 0.2 kW/cm2 was periodically turned
on and off, the fluorescence signal was monitored by confocal
microscopy at room temperature. As expected, the fluorescence
intensity of GFP decreased slowly and did not recover in
subsequent pulses. It is well-known that GFP’s signal drop
comes from irreversible photobleaching18 and/or photo-
conversion to a long-lived dark state.19
In contrast, when the same excitation scheme was applied to
Spinach in the presence of excess DFHBI (5 μM), the initially
bright fluorescence of the Spinach−DFHBI complex showed
fast decay to a significantly reduced fluorescence level (<5% of
the initial level) within 2 s. Remarkably, after the excitation light
was switched off for a few seconds, a subsequent pulse revealed
that the fluorescence signal was recovered back to 85−95% of
the initial signal. This indicates that the fluorescence loss of
Spinach−DFHBI is not due to fast, irreversible photobleaching
but instead is caused by reversible conversion to a non-
fluorescent state and that the signal recovery occurs
spontaneously without light.
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Figure 2. (a) Fluorescence time traces and (b) decay rates of
immobilized GFP and Spinach−DFHBI complex as a function of Iexc
.
Figure 3. Time-resolved fluorescence of the Spinach−DFHBI
complex. The fluorescence lifetime was measured when the degree
of photoconversion was maximized (Spinach CW, top left) or
minimized (Spinach pulsed, top right). However, both sets of
conditions resulted in the same decay rate, τf = 4.0 ns. For
comparison, the decay curves of GFP (green line; τf = 2.7 ns) and
DFHBI-only solution (gray dotted line; τf < 80 ps) are shown.
The fluorescence intensity is normalized to the maximum value at the
highest Iexc, and the decay rate (koff) is the inverse of the half-life.
[DFHBI] = 5 μM. (c) Fluorescence intensity time traces of Spinach
free in solution (solid lines). The red dotted line is for GFP free in
solution at Iexc = 160 kW/cm2.
Power-Dependent Fluorescence Decay. We next
examined how the excitation intensity Iexc affects the
fluorescence decay (Figure 2a). The fluorescence decay of
GFP becomes faster at higher Iexc, and its decay rate (koff)
increases linearly with Iexc (Figure 2b). For Spinach, however,
used 0.5 μM Spinach and 0.5 μM DFHBI free in solution. As
depicted in Figure 3, in both cases (i.e., regardless of the extent
of photoconversion), the complex exhibited a single-exponen-
tial decay with τf = 4.0 0.1 ns, which is 1.5 times longer than
that of GFP (τf = 2.7 ns). This suggests that the photo-
converted species does not accumulate to a high degree to be
detected based on the fluorescence lifetime. The concentration
of DFHBI does not greatly affect the fluorescence lifetime of
the Spinach−DFHBI complex (Figure S2 in the SI).
koff exhibits a weaker dependence on Iexc: it changes only 2-fold
(from 11.7 to 21.2 s−1) when Iexc increases 25-fold from 0.2 to 5
kW/cm2, a range of Iexc commonly used in confocal microscopy
and single-molecule imaging (Figure 2b). The very weak
dependence of the fluorescence decay rate on Iexc suggests that
fluorescence loss requires at least two steps, an initial light-
dependent step already saturated at our typical Iexc values and a
second light-independent step.
[DFHBI]-Dependent Fluorescence Recovery. These
characteristic behaviors of the Spinach−DFHBI complex may
be attributed to (i) reversible photoconversion as previously
proposed,20 (ii) fluorogen exchange,11 or (iii) a combination of
the two. To further clarify the mechanism, we next determined
the fluorescence recovery rate as a function of fluorogen
concentration. If the Spinach−DFHBI complex is photo-
converted to a nonfluorescent state in which the fluorogen is
still bound to the RNA aptamer, thermal or light-induced
recovery to a fluorescent state would be independent of
[DFHBI]. Instead, the recovery rate (kon) displays a strong
dependence on [DFHBI]: kon changes from 0.08 to 2.0 s−1 as
[DFHBI] increases from 1 to 200 μM (Figure 4). This result is
in disagreement with a previous proposal20 where the reported
recovery rate was constant at ∼0.0027 s−1 independent of the
DFHBI concentration. This discrepancy results from the fact
that we used immobilized RNA in an excess of DFHBI, whereas
in the latter case, the fluorescence recovery measured the
replenishment rate of the fluorescent Spinach−DFHBI complex
in a freely diffusing solution (please see the SI for detailed
discussion). The sublinear behavior of the recovery rate in
Figure 4b might be attributed to altered spectral properties of
the Spinach−DFHBI complex at high concentrations of
DFHBI, although its molecular mechanism remains unclear
(Figure S3 in the SI).
For the free species in solution, the fluorescence decay would
be difficult to detect because of replenishment of fluorescently
active species from outside the excitation volume via diffusion.
This was indeed the case for GFP (Figure 2c). For Spinach−
DFHBI, however, photoconversion is very efficient, saturating
even at moderate Iexc values (Figure 2a), and as a result, we
could observe significant fluorescence decays even without
immobilization (Figure 2c). To achieve the same degree of
fluorescence decay for GFP, we had to use 1000 times higher
Iexc (Figure 2c and Figure S1 in the SI). This set of
measurements on free species in solution further indicates
that the weak Iexc dependence of the fluorescence decay kinetics
of immobilized Spinach−DFHBI is due to saturation of the
highly efficient photoconversion process (please see the SI for
further discussion).
Fluorescence Lifetime of the Spinach−DFHBI Com-
plex. We examined the time-resolved fluorescence of the
Spinach−DFHBI complex in two different ways to gain further
insights. First, the fluorescence lifetime (τf) was measured when
>80% of the complex was photoconverted after 8 s of
illumination (Figure 3, top left). Second, τf was measured
during a 20 ms pulse of illumination time every 2 s to ensure
collection of the fluorescence mainly from the bright molecular
complex before photoconversion (Figure 3, top right). Here we
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continuous-wave (CW) illumination or under pulsed illumina-
tion (50 ms pulse width) at a repetition rate varying from 2 to
0.2 Hz (Figure 6a). A similar scheme has been applied in
fluorescence microscopy to attain high fluorescence yields from
organic dyes and fluorescent proteins, where the repetition
rates ranged from 0.1 to 100 kHz to allow dark-state
relaxation.22
To demonstrate the concept, we transformed E. coli with
plasmids expressing Spinach and imaged them in the presence
of 100 μM DFHBI (Figure 6b).11 Time-lapse fluorescence
microscopy showed that the fluorescence signal of E. coli under
CW illumination rapidly drops to <10% of the initial intensity
within 0.2 s (Figure 6c), whereas pulsed illumination at a
repetition rate of 0.2 Hz retains the fluorescence signal much
better by ensuring sufficient recovery time between successive
pulses (Figure 6d). This phenomenon holds true for
immobilized Spinach RNA aptamers as well (Figure S4 in the
SI). We further quantified the data by plotting the fluorescence
time trajectories of a single cell (Figure 6e) and 40 cells
averaged (Figure 6f).
Figure 4. (a) [DFHBI]-dependent fluorescence time traces and (b)
recovery rates of the Spinach−DFHBI complex. The inset in (b)
shows the recovery rate at low [DFHBI] with a linear fit (red line).
The accumulated fluorescence intensity, representing the
total photon flux, is 5.7-fold higher at 0.2 Hz than with CW
illumination for the same 2 s of total illumination time (Figure
6g). The steady-state intensity is about 10-fold higher at 0.2 Hz
compared with CW illumination (Figure 6e,f). Although the
lower repetition rate gives higher fluorescence signal gain
(Figure 6h), there is a trade-off in imaging speed. Different
RNA aptamers selected for higher kbind may allow even higher
imaging speed. Additionally, pulsed illumination with longer
pulse widths is not effective in gaining higher signal and
requires more recovery time, probably because shorter pulses
(10−50 ms) minimize the photoconversion process (Figure
6i).
Photophysical Model of the Spinach−DFHBI Com-
plex. On the basis of our results, we can rule out the
possibilities that Spinach undergoes reversible photoconversion
without fluorogen exchange [model (i)] or efficient fluorogen
exchange without photoconversion [model (ii)]. A study of a
malachite green RNA aptamer has shown that the fluorogen
can go through significant changes in its conformation and
electronic structure upon illumination.23 In the case of
photoswitchable proteins, photoisomerization is known to be
strongly involved with the formation of the dark state,24 where
an accompanying change in the configuration of fluorescent
proteins leads to the conversion of a (fluorescent) deproto-
nated state into a (nonfluorescent) protonated state.25 We
confirmed that after 405 nm illumination the absorbance of free
DFHBI decreases, with its spectrum showing a red-shifted band
and an enhanced high-energy band (Figure S5a in the SI). This
is very similar to the spectroscopic features of GFP
chromophore and its analogues, indicating that cis/trans
photoisomerization occurs for DFHBI.26 For these reasons, it
is tempting to suggest that the photoconversion of the
Spinach−DFHBI complex may also be related to cis/trans
photoisomerization, although a definite conclusion should await
further studies.20,27
Figure 5. Binding−unbinding kinetics of the Spinach−DFHBI
complex. (a) Fluorescence−time profile when 40 nM Spinach was
mixed with 200 nM DFHBI under a very low light level. The red line is
a monoexponential fit. (b) Determination of the binding (kbind) and
unbinding (kunbind) kinetic rates from the linear fit (red solid line) of
kobs vs concentration.
Binding−Unbinding Kinetics. Next we examined how fast
fluorogen exchange occurs in the Spinach−DFHBI complex.
Spinach RNA was mixed with DFHBI in a quartz cuvette, and
the fluorescence change was monitored at a very low light level
(<1 to 5 W/cm2) to minimize complications arising from
photoconversion. Figure 5a shows that the fluorescence signal
rises monoexponentially, giving the observed rate constant
(kobs). The slope of the linear fit of the plot of kobs versus the
concentration of Spinach and DFHBI (Figure 5b) directly
provides the binding rate constant, kbind, and intercept gives the
unbinding rate constant, kunbind. The obtained values, kbind
=
(6.2 0.1) × 104 M−1 s−1 and kunbind = (2.4 0.1) × 10−2 s−1,
yield a dissociation equilibrium constant KD = 390 nM, which is
in good agreement with previous studies.11,20 Here, kbind of
Spinach−DFHBI is well within the range of typical small-
molecule−RNA aptamer interactions, which vary from 103 to
106 M−1 s−1.21
It should be noted that the dissociation time (1/kunbind) is
very long, and therefore, once Spinach binds to DFHBI, it stays
as a complex for >40 s under a low light level or in the absence
of light. Also, the [DFHBI] dependence of the recovery rate at
low concentrations gives a bimolecular reaction rate of (4.9
0.3) × 104 M−1 s−1, which corresponds well to kbind (Figure 4b
inset). This confirms that the fluorescence recovery is mainly
due to a bimolecular reaction between Spinach and DFHBI
rather than a spontaneous recovery of the photoconverted
complex.
Fluorescence Signal Gain with Low-Repetition-Rate
Illumination and Its Application to E. coli Imaging. The
limited time duration of the fluorescence of the Spinach−
DFHBI can be mitigated by low-repetition-rate illumination.
Here the fluorescence intensity was monitored under
To account for the unique photophysical behavior of
Spinach, we propose the alternative model shown in Figure
7. Upon illumination of the Spinach−DFHBI complex, the
bound DFHBI undergoes photoisomerization at a high kinetic
rate (ko*ff). It is possible that the photoisomerized complex
before unbinding is already completely dark. However, because
of the lifetime measurements shown in Figure 3, we can rule
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Figure 6. Fluorescence signal gain from living E. coli expressing Spinach with low-repetition-rate illumination. (a) Scheme of illumination using CW
and pulsed modes. (b) Spinach and DFHBI are necessary for imaging of bright fluorescence from E. coli. Scale bar = 5 μm. (c, d) Time-lapse
fluorescence images of E. coli under (c) CW and (d) pulsed illumination with the same peak intensity (∼100 W/cm2). [DFHBI] = 100 μM; scale bar
= 5 μm. (e) Fluorescence intensity traces from the single cell marked by white arrows in (c) and (d). (f) Repetition-rate-dependent fluorescence
intensity traces averaged over many cells (n > 40). (g) Cumulative fluorescence intensity at various repetition rates. (h) Cumulative fluorescence
intensity after 2 s of total illumination time. The CW data point is marked by a black square dot. Inset: Summed images of a single E. coli (scale bar =
1 μm). (i) Longer pulse illumination (Ton = 5 s) leads to less signal gain and longer recovery time.
out the possibility that a partially quenched photoisomerized
complex accumulates significantly under our experimental
conditions. Photoisomerization results in fast unbinding
(ku*nbind) of the photoisomerized fluorogen, in contrast to the
long-lived binding of DFHBI under very low or no illumination.
Such accelerated unbinding could be caused by two factors:
first, since the RNA aptamer was selected on the basis of its
binding affinity to the ground-state of DFHBI, its binding
affinity to a photoisomerized DFHBI may not be optimal.
Second, the Spinach aptamer might also change its con-
formation to accommodate photoisomerized DFHBI, a
phenomenon observed in photoconversion of fluorescent
proteins, which could greatly decrease or even abolish binding.
Rapid dissociation of photoisomerized DFHBI allows Spinach
to bind to a new ground-state DFHBI from solution, restoring
the fluorescence. This could also explain why we could not
identify the photoisomerized product via fluorescence lifetime
Figure 7. Proposed model of the fluorescence behavior of the
Spinach−DFHBI complex. The two isomers of DFHBI are depicted at
the bottom.
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CONCLUSION
■
It was reported that the basic fluorescence properties of the
Spinach−DFHBI complex are similar to those of enhanced
GFP.11 This seems plausible given the similar ways in which the
scaffolds of GFP and the Spinach RNA aptamer make a
nonfluorescent chromophore fluorescent through structural
reinforcement involving specific hydrogen-bonding and π-
stacking interactions. However, our results show that the
different nature of the RNA aptamer endows unique properties,
namely, fast fluorescence decay and [DFHBI]-dependent
recovery attributed to photoconversion accompanied by
accelerated unbinding. The use of pulsed illumination greatly
increases the steady-state fluorescence intensity and total
photon flux of the Spinach−DFHBI complex. Combined with
signal amplification by attachment of many repeats of the RNA
aptamer to a target RNA,29 this pulsed excitation scheme may
allow cellular RNA imaging at the single-RNA level with high
signal-to-noise ratio. Our study may also provide insights into
how to improve the fluorescence properties of the Spinach−
DFHBI complex as well as various fluorogenic dye−RNA
aptamer complexes.29,30
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W.; Jakobs, S. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 13005.
(25) Tolbert, L. M.; Baldridge, A.; Kowalik, J.; Solntsev, K. M. Acc.
Chem. Res. 2012, 45, 171. Dedecker, P.; De Schryver, F. C.; Hofkens, J.
J. Am. Chem. Soc. 2013, 135, 2387.
ASSOCIATED CONTENT
* Supporting Information
Experimental details, notes on fluorescence decay in a freely
diffusing solution and on the measurement of recovery rates,
and Figures S1−S5. This material is available free of charge via
the Internet at http://pubs.acs.org.
■
S
AUTHOR INFORMATION
Corresponding Author
tjha@illinois.edu
■
Notes
(26) Voliani, V.; Bizzarri, R.; Nifosi, R.; Abbruzzetti, S.; Grandi, E.;
Viappiani, C.; Beltram, F. J. Phys. Chem. B 2008, 112, 10714.
(27) Addison, K.; Conyard, J.; Dixon, T.; Page, P. C. B.; Solntsev, K.
M.; Meech, S. R. J. Phys. Chem. Lett. 2012, 3, 2298.
(28) Strack, R. L.; Disney, M. D.; Jaffrey, S. R. Nat. Methods 2013, 10,
1219.
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank Dr. Sung Chul Bae for providing the 473 nm laser
and Prof. Samie R. Jaffrey for providing the pET28c-tRNA-
Spinach plasmid. Funding was provided by NIH (GM065367
and AI083025) and NSF (PHY0822613). T.H. is an
investigator of the Howard Hughes Medical Institute.
(29) Babendure, J. R.; Adams, S. R.; Tsien, R. Y. J. Am. Chem. Soc.
2003, 125, 14716.
(30) Sparano, B. A.; Koide, K. J. Am. Chem. Soc. 2007, 129, 4785.
Constantin, T. P.; Silva, G. L.; Robertson, K. L.; Hamilton, T. P.;
Fague, K.; Waggoner, A. S.; Armitage, B. A. Org. Lett. 2008, 10, 1561.
Sando, S.; Narita, A.; Hayami, M.; Aoyama, Y. Chem. Commun. 2008,
3858. Lee, J.; Lee, K. H.; Jeon, J.; Dragulescu-Andrasi, A.; Xiao, F.;
Rao, J. ACS Chem. Biol. 2010, 5, 1065.
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